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Histone chaperones and the Rrm3p helicase regulate flocculation in S. cerevisiae
Epigenetics & Chromatin volume 12, Article number: 56 (2019)
Biofilm formation or flocculation is a major phenotype in wild type budding yeasts but rarely seen in laboratory yeast strains. Here, we analysed flocculation phenotypes and the expression of FLO genes in laboratory strains with various genetic backgrounds.
We show that mutations in histone chaperones, the helicase RRM3 and the Histone Deacetylase HDA1 de-repress the FLO genes and partially reconstitute flocculation. We demonstrate that the loss of repression correlates to elevated expression of several FLO genes, to increased acetylation of histones at the promoter of FLO1 and to variegated expression of FLO11. We show that these effects are related to the activity of CAF-1 at the replication forks. We also demonstrate that nitrogen starvation or inhibition of histone deacetylases do not produce flocculation in W303 and BY4742 strains but do so in strains compromised for chromatin maintenance. Finally, we correlate the de-repression of FLO genes to the loss of silencing at the subtelomeric and mating type gene loci.
We conclude that the deregulation of chromatin maintenance and transmission is sufficient to reconstitute flocculation in laboratory yeast strains. Consequently, we propose that a gain in epigenetic silencing is a major contributing factor for the loss of flocculation phenotypes in these strains. We suggest that flocculation in yeasts provides an excellent model for addressing the challenging issue of how epigenetic mechanisms contribute to evolution.
The non-sexual aggregation of single cell organisms into clusters is referred to as flocculation or biofilm [1, 2]. In industrial yeast strains flocculation is a highly desired phenotype and in many cases can be readily activated by starvation, exposure to ethanol and/or other stressors [1, 2].
The key regulators of flocculation in S. cerevisiae are the FLO genes. They are positioned 20–40 kb away from the telomeres and encode lectin-like cell surface proteins [3, 4]. The genes contain multiple internal repeats and share significant homology with FLO genes in other yeast species [3, 5]. FLO1 acts as a regulator of biofilm formation  while FLO11 is known to control the switch between planktonic and filamentous growth . Other members of the family include FLO5 (paralogous to FLO1), FLO9 and FLO10 . In lab strains the FLO genes are repressed by the Tup1/Cyc8 complex via long-range chromatin remodelling . FLO11 is reversibly switching between active and silent states, a feature reminiscent of subtelomeric genes [4, 8].
FLO expression and flocculation is regulated by a wide variety of mechanisms including the MAPK, TORC, SNF1 and RIM101 signalling cascades [9, 10]. Chromatin structure plays a major role in the regulation of flocculation, but details are often missing. For example, screens in the ∑1278b strain (unlike S288C, ∑1278b displays various dimorphic transitions) have shown that flocculation and filamentous growth are suppressed by mutations in components of the histone deacetylase Rpd3, the acetyl-transferase SAGA or the Ino80/Swr1p chromatin remodeler [9, 10]. In industrial yeasts, the Set1/COMPASS histone methyl transferase and the RPD3, HDA1 and HST1 deacetylases have been implicated in the repression of FLO genes [4, 7, 11]. Finally, a mutation in Histone H4 (H4S47C) leads to the depression of FLO1 and flocculation . Importantly, the major regulators of gene silencing in S. cerevisiae, the SIR genes, have not been pulled out in any of these screens. Instead, it has been shown that in laboratory strains the FLO genes are repressed by the HST1 and HST2 paralogs of the SIR2 histone deacetylase . Even more, in wine yeasts SIR2 is required for the expression of FLO11 while the acetyl transferase SAS2 represses the transcription of FLO5 .
Several histone chaperones are involved in the epigenetic transmission and maintenance chromatin structure in S. cerevisiae [14, 15]. The histone chaperones ASF1 and FACT are involved in both the disassembly and reassembly of nucleosomes during DNA replication. CAF-I is believed to play a central role in the re-assembly of H3/H4 tetramers behind the forks while ASF1 and Rtt106 participate in the delivery of new H3 and H4 histones . ASF1 and FACT also have roles in transcription that is independent of their function in DNA replication . On the other hand, the HIR and NAP1 chaperones operate in a replication-independent manner, but their precise role is not clear . No reports have linked the repression of FLO genes to histone chaperones. However, there is solid evidence for replication-coupled chromatin assembly factors contributing to gene silencing at the sub-telomeres and the mating type HMR/HML loci [14, 16,17,18,19]. In addition, we have shown that Rrm3p, a DNA helicase that removes tightly bound proteins ahead of the replication forks, has a role in the mechanism of epigenetic conversions at the sub-telomere .
Interestingly, many laboratory S. cerevisiae strains contain functional copies of the FLO genes but do not normally flocculate, most likely because of the extensive passive selection against flocculation in favor of planktonic growth [1, 20]. We have recently noticed that mutations in various histone chaperones promote flocculation-like phenotypes. In this manuscript, we report our extensive analyses of these observations.
Flocculation-like phenotypes in laboratory strains
While analysing epistatic interactions of histone chaperones with the RRM3 helicase, we noticed that some mutant strains produced clusters in liquid cultures. This was surprising as all mutations were in haploid BY4742 or W303 backgrounds, which do not flocculate under normal laboratory conditions. We systematically compared these and other phenotypes of all strains listed Table 1 plus about another 120 strains with mutations in various genes (not shown).
No flocculation-like phenotypes were observed in any of the single deletion mutants in BY4742 and W303 genetic background, thus reiterating the notion that these laboratory strains have lost the ability to form biofilm (Fig. 1a). On the other hand, when liquid cultures were grown on a spinning wheel visible clusters of cells were observed in strains with some, but not all combinations of double deletions of cac1, asf1, hir1 and rrm3. As already mentioned, CAC1, ASF1 and HIR1 encode for histone chaperones engaged in the assembly of nucleosomes  while RRM3 encodes a helicase that relieves replication pausing . Flocculation was apparent as visible clusters in liquid cultures as compared to the uniform turbid appearance of the non-flocculating strains (Fig. 1a). Even more, the cultures formed pellets shortly after removal from the wheel (Fig. 1b). The levels of sedimentation of these laboratory strains were comparable to the sedimentation of a wild type beer strain with a well documented flocculation phenotype (Hornindal 1 ) (Fig. 1b). In all cases, these clusters were dispersed by exposure to 10 mM EDTA (not shown) as previously observed in many wild type strains .
It has been reported that in feral and industrial strains flocculation is often accompanied by altered colony morphology, stronger adhesion to agar and cell cycle arrest . However, we noticed that our flocculating strains formed even lustre colonies similar to the ones formed by non-flocculating strains (Fig. 1c) with no evidence of stronger adhesion to agar as compared to the isogenic wild type strains (not shown). We also found no correlation between growth rates of the strains, their progression through the cell cycle (Additional file 1) and flocculation (Fig. 1a). Even more, individual isolates of some strains displayed various levels of flocculation as judged by the observation of clusters under a microscope (not shown) while growth rates were the same.
We concluded that double deletion mutants in the BY4742 and W303 laboratory strains promote apparent cell aggregation, but not a full display of other flocculation-related phenotypes observed in some wild type S. cerevisiae strains.
Recombination of FLO genes, spontaneous mutations and sensitivity to DNA damage do not correlate to flocculation-like phenotypes
Prior studies have reported variations in the number of intragenic repeats of FLO genes of industrial yeasts and have suggested that homologous recombination and length variations could have phenotypic and evolutionary implications [5, 24]. In addition, it has been previously reported that cac1∆ and rrm3∆ strains have elevated spontaneous mutation rates and sensitivity to DNA damage [16, 17, 25]. For this reason, we analysed the length of FLO genes in our laboratory strains as in . We also looked at the frequency of spontaneous mutations as measured by the canavanine resistance fluctuation assay  and at the sensitivity to DNA damage as measured by exposure to Methyl Methane Sulfonate (MMS). We observed length variation only in FLO10 in the cac1∆asf1∆ strain, but not in any of the FLO genes in any of the flocculating laboratory strains (Additional file 1: Figure S3). The canavanine resistance assay measures the rate of spontaneous loss-of-function mutations in the CAN1 arginine transporter gene and was performed only in the strains that do not already harbor the can1-100 mutation. These limited in scope assays indicated a modest increase in spontaneous mutation rates in rrm3∆tof1∆ and cac1∆tof1∆ which do not flocculate. In all other tested strains the mutation rates were indistinguishable from the wild type BY4742 strain (Additional file 1). Finally, there was no correlation between sensitivity to MMS (Additional file 1) and flocculation of the strains.
We concluded that the flocculation in our laboratory strains is not related to any of these earlier reported phenotypes and characteristics in various single deletion mutants.
Elevated expression of FLO genes in flocculating strains
In industrial yeasts, the FLO genes are repressed in planktonic cultures and active in flocculating ones [4, 6]. We asked if the flocculation phenotype in our strains could be attributed to the elevated expression of the FLO genes. RNA was isolated from four flocculating and four non-flocculating strains and analysed by qRT-PCR with primers specific for each of FLO1, FLO9, FLO10 and FLO11. Because of the highly repetitive nature of the high degree of homology between the FLO genes, the primers for FLO5 also amplify the RNAs produced by FLO1 and FLO9. Three to five independent experiments were performed with each strain/primer combination and the measured amounts in the mutants were compared to the expression of the FLO genes in the isogenic BY4742 strain. The analyses showed 1- to 2-fold increase in the expression of FLO1 and FLO9 and 2- to 3-fold increase in FLO10 and FLO11, respectively, in the non-flocculating single deletion mutants cac1∆, asf1∆, rrm3∆ and hir1∆ (Fig. 2). On the other hand, the expression of all FLO genes was 4- to 12-fold higher in the flocculating double deletion mutants. The FLO1/FLO5/FLO9 primers detected 10- to 80-fold increase in the abundance of these RNAs (Fig. 2). While the overexpression of FLO genes in flocculating strains was consistently observed, the magnitude of effects differed between individual experiments (Fig. 2). While we cannot confidently explain these fluctuations, we suspect that FLO gene expression varies in individual cultures and that this variation could be an important component in the adaptation/flocculation strategy of the cells. This notion is consistent with the observed difference of flocculation between individual isolates of the same strains, as mentioned above. Regardless of the nature of these variations, we observed a consistent correlation between the flocculation phenotype and the higher expression of the FLO genes.
Hyper-acetylation of histones H3 and H4 at the FLO loci in the flocculating strains
Earlier studies have shown that the repression of FLO1 is dependent on the HST1, HST2, HDA1 and RPD3 histone deacetylases and that FLO1 de-repression is associated with the hyperacetylation of Histones H3 and H4 at its promoter [7, 26]. While the mutants we have analysed so far do not encode histone deacetylases, we reasoned that their effects could nevertheless be mediated by hyperacetylation of Histones H3 and H4 at the FLO genes promoters. We addressed this question by Chromatin-Immuno-Precipitation (ChIP) with anti-H3, anti-H3AC and anti-H4AC antibodies followed by quantitative PCR with primers for the promoter regions of FLO1 and FLO11. Consistent with earlier observations, at the FLO1 promoter the flocculating strains produced 2–11 times higher signal with the anti-H3AC antibody and 3–6 times higher signal anti-H4AC antibodies, as compared to the BY4742 strain (Fig. 3a). On the other hand, there was no difference in the acetylation of histones at the FLO11 promoter between the flocculating strains and BY4742 (Fig. 3b). We suspect that we were not able to detect difference in H3/H4 acetylation at the FLO11 promoter because FLO11 switches between active and silent states  thus producing a higher basal signal in ChIP experiments. It has also been shown that FLO1 remains active while FLO11 is repressed depending on the abundance of Tup1p and Cyc8p . Additionally, it remains possible that our RT-qPCR (see above) and GFP expression driven by the FLO11 promoter (see below) analyses are sensitive enough to reveal transient increases in FLO11 expression but not temporary acetylation changes by ChIP assays.
We also quantified the H3-ChIP signals at ACT1 and the FLO1, FLO11 promoters. The results showed that in the cac1∆asf1∆, asf1∆rrm3∆ and cac1∆hir1∆ strains the H3-ChIP signals decline relative to W303 at these three loci with a similar but less pronounced effect in cac1∆rrm3∆ (Fig. 3c). We also tested the sensitivity of the chromatin in these strains to micrococcal nuclease (MNase) (Additional file 1: Figure S7). In agreement with the H3-ChIP data, we observed substantially higher sensitivity to MNase in cac1∆asf1∆, asf1∆rrm3∆ and cac1∆hir1∆ cells and modest increase in cac1∆rrm3∆ cells relative to W303. These results are consistent with earlier observations showing increased sensitivity to nucleases in cac1∆ and decreased sensitivity in asf1∆ . However, the deletion of additional genes in our strains seems to exacerbate the altered sensitivity to nucleases and lead to a profound de-repression of the FLO genes.
Taken together, the ChIP, MNase sensitivity and RT-PCR data point to a lower nucleosome density and higher H3/H4 acetylation at the FLO promoters that contributes to the loss of repression of the FLO genes.
Epistatic interactions of CAC1, ASF1 and HIR1 with HDA1
We reasoned that if histone chaperones are indeed playing a central role in the transmission and maintenance of repressive chromatin at the FLO loci, then CAC1, ASF1 and HIR1 would genetically interact the histone deacetylases HDA1 or RPD3 to promote flocculation phenotypes. We constructed haploid cac1∆hda1∆, asf1∆hda1∆ and hir1∆hda1∆ strains, but were not successful in producing double deletion mutants with RPD3 with any of these genes. We observed apparent flocculation in the cac1∆hda1∆ and asf1∆hda1∆ strains, but not in the hir1∆hda1∆ strain (Fig. 4). Similarly, no flocculation was observed in the hir1∆rrm3∆ strain. These observations suggest that HIR1 is not normally involved at the replication forks, but might have a role in FLO gene repression by yet unknown mechanism.
Flocculation in single deletion mutants is induced by nitrogen starvation or by inhibition of histone deacetylases
Nicotinamide (NAM) is a potent competitive inhibitor of NADH+-dependent histone-deacetylases . Cells exposed to NAM display reduced silencing of subtelomeric genes and at the mating type loci , however, its effect on flocculation has not been reported. We tested this possibility by growing liquid cultures of various strains in the presence of 0, 2 and 5 mM NAM. The cultures were then rested for 30 min and the rate of sedimentation in the NAM-treated relative to the non-treated samples was measured. Three cultures per strain were scored on 3 different days. Flocculation was further confirmed by light microscopy as in Fig. 1. Sedimentation scores are shown in Additional file 1: Table S2 and the results are summarized in Fig. 5a. In the presence of 5 mM NAM we observed 3–4 times faster sedimentation rates in cac1∆ and asf1∆ strains and 2 times faster sedimentation rates in hir1∆ and rtt106∆ strains in both W303 and BY4742 background (Fig. 5a). The rrm3∆ strain showed less than twofold faster rates. All other strains tested, including BY4742, W303, hst1∆, tof1∆, sir2∆, gcn5∆ (Fig. 5a) or the already flocculating double deletion mutants (not shown) revealed no apparent detectable increase in sedimentation rates in the presence of NAM.
So far we have shown that strains with single deletions do not produce floccules but do so in combination with the deletion of other genes or upon incubation with NAM (Figs. 1, 5a). On the other hand, it has been reported that stress and starvation induce flocculation in wild type yeast strains . We asked if the single deletion mutants would produce floccules in media containing low nitrogen or low carbon source. The low carbon medium had little effect on the flocculation of any of these strains (not shown). On the other hand, upon nitrogen starvation we readily observed flocculation in the cac1∆, asf1∆ and hir1∆, but not in the isogenic BY4742 strain or in the rrm3∆ strain (Fig. 5b).
These experiments indicated that the repression of the FLO genes is already compromised in cac1∆, asf1∆, hir1∆ and rtt106∆ mutants and can be further revealed by nitrogen starvation or treatment with NAM.
Loss of the association of CAF-1 with PCNA induces flocculation
We asked if our observations depend on the activity of CAF-1 in a replication-dependent or independent fashion. We used two strains that harbor mutations in the replication clamp PCNA (pol30-6 and pol30-79), which are known to significantly reduce its association with CAF-1 . We reasoned that these PCNA mutations could produce effects similar to these seen in cac1∆ mutants. As expected, the deletion of RRM3 and ASF1 in the pol30-6 and pol30-79, but not POL30 strains produced apparent flocculation (Fig. 6a). Interestingly, the deletion of RRM3 produced a stronger effect in the pol30-6 mutant and the deletion of ASF1 produced a stronger effect in the pol30-79 mutant (Fig. 6a). At present, we cannot explain the differences between the two pol30 alleles. Next, we complemented the cac1∆hir1∆, cac1∆rrm3∆ and cac1∆asf1∆ strains with CAC1 or cac1∆PIP with a destroyed PCNA Interacting Peptide (PIP), which is known to preclude the association of CAF-1 with PCNA and CAF-1 mediated replication-coupled chromatin assembly in vitro . In Fig. 6b we show that complementation by CAC1 substantially reversed their flocculation phenotype while cac1∆PIP had no effect. Finally, we complemented a cac1∆ strain with CAC1 and cac1∆PIP and exposed them to Nicotinamide (NAM). In Fig. 6c we show that NAM-induced rates of sedimentation, similar to what is observed in cac1∆1 mutants in the strain complemented with cac1∆PIP but not by CAC1. Together, these experiments demonstrated that the inability of CAF-1 to associate with PCNA can induce flocculation.
Variegation and loss of FLO11 silencing in a flocculating strain
It has been previously demonstrated that a wild type yeast strain expresses FLO11 in a variegated fashion producing patches of GFP+ and GFP− cells when the gene is tagged with GFP  and that FLO11 is contributing to filamentous growth . We, therefore, asked if flocculation can be correlated to the variegated expression of FLO11. We replaced the FLO11 ORF with GFP as in  and tested the expression pattern of GFP in a flocculating (cac1∆rrm3∆) and two non-flocculating laboratory (cac1∆ or rrm3∆) strains in W303 genetic background. Briefly, cells were dispersed, serially diluted in 96-well trays and incubated without shaking for 24 h. Sparsely populated wells with isolated cell clusters were then analysed by fluorescent microscopy. In cac1∆, rrm3∆ and the control W303 strains we observed a very low number of isolated GFP+ cells (Fig. 7a). In contrast, the flocculating cac1∆rrm3∆ displayed clusters of cells with various levels of green fluorescence as well as cells with no apparent fluorescence (Fig. 7a).
Because cell clusters do not allow for a precise focusing and measurement of the GFP signals, the strains were grown in suspension for 24 h, dispersed by vigorous vortexing/pipetting and spread on microscope slides. GFP signals were acquired for both individual GFP+ and GFP− cells and used for the measurement of GFP+ signals in individual cells and for the calculation of the percent of GFP+ cells in each strain. In agreement with the observations in cell clusters (Fig. 7a), these measurements showed that the signals from individual cac1∆rrm3∆ cells significantly vary, but on average were substantially higher than the signals in cac1∆, rrm3∆ or the control W303 strains (Fig. 7b). We suspect that the difference in the detected levels of FLO11 RNA (Fig. 2) and GFP (Fig. 7) could be caused by the stability of GFP. Next, the percent of GFP+ cells was calculated. GFP+ cells were defined as cells that have 3 times higher green fluorescence as compared to the average fluorescence from the GFP− cells. Based on these criteria, we show about 45% GFP+ cells in the cac1∆rrm3∆ strain as compared to 8%, 9% and 4% in the cac1∆, rrm3∆ and W303 strains, respectively.
We concluded that flocculation in the cac1∆rrm3∆ strain is accompanied by a gain of FLO11 gene expression in about half of the cells in the culture. We also concluded that the expression of GFP as driven by the FLO11 promoter varies between individual cells. Finally, because we did not observe large patches of GFP+ and GFP− cells in the cell clusters (Fig. 7a, left panel), we suspect that the conversion rates between active and silent FLO11 are high and cannot be assessed by the methods we have used in the past for the analysis of conversion rates at the telomeres .
Correlation of repression of FLO genes and the sub-telomeric and the mating type loci in laboratory yeast strains
Next, we looked at data from earlier publications by us and others [4, 16, 19, 25, 34] to compare flocculation phenotypes and FLO gene silencing to the level of gene silencing at the telomeres and the mating type loci (Table 1). Consistent with earlier observations [4, 34] and this study, SIR3 was required only for gene silencing at the sub-telomeres and the mating type loci and had no effect on the expression of the FLO genes. At the same time, the single and double deletions of histone chaperones and RRM3 seemed to have a similar magnitude of effect on all these loci. For example, single deletions of CAC1, ASF1, HIR1 and RRM3 do not or only transiently de-repress the mating type loci [16, 19, 25, 34], moderately reduce the repression of FLO genes (this study) and moderately reduce gene silencing at the telomeres [16, 25]. In comparison, double deletions of these genes cause a measurable loss of silencing at the mating type loci ([19, 25, 34] and Additional file 1: Figure S6), significant de-repression of the FLO genes (this study) and severe loss of silencing at the telomeres [16, 25].
Flocculation phenotype in an ancestral strain
Next, we asked if an ancestral strain (EM93, ) used to produce S288C and eventually BY4742 and W303 is flocculating under normal laboratory conditions. We obtained an old stock of EM93, which was heavily flocculent. We produced four independent clones form dissected individual spores and grew them for at least 60 generations. Because the EM93 strain is HO+ and is capable of switching its mating type, by the time of the assessment of flocculation these cultures have become diploid as confirmed by test growth in sporulation medium and observation of tetrads. All EM93 clones displayed heavy flocculation (Fig. 8). Next, we disrupted HO and produced four haploid EM93 clones, which also displayed flocculation, but not as strong as the diploid clones. These observations indicated that the original feral strain had the ability to flocculate in both haploid and diploid state, however, this phenotype was lost through the subsequent breeding and selection for planktonic growth .
Here, we report the partial reconstitution of flocculation in laboratory yeast strains, which have lost this phenotype by continuous passive selection for planktonic growth . The precise reasons for the loss of flocculation are not clear. It has been reported that in the W303 and BY4742 strains FLO8 (a transcription factor for the FLO genes) harbors an inactivating point mutation , while variation in FLO11 were correlated to the phenotype of Σ1278b . We show that, in addition to this and possibly other gene mutations, the loss of flocculation is due to a significant gain in FLO gene silencing by epigenetic means. In support, we tested 120 mutant laboratory strains (not shown) and demonstrate that flocculation phenotypes can be reconstituted by the deletion of histone chaperones, histone deacetylases and a DNA helicase that relieves replication pausing (Fig. 1, Additional file 1: Table S2 and data not shown). We also show that the EM93 strain, which is ancestral to S288C and W303, has apparent flocculation phenotype when grown under normal laboratory conditions. These experiments do not reveal exactly how the feral strain has lost its flocculation trait. In comparison, a continuous culturing and analysis of other feral strains has shown their ability to produce various colony morphologies and flocculation phenotypes . It is quite possible that a similar process of adaptation/domestication has produced the non-flocculating phenotype of W303. Our study suggest that epigenetics could be an important factor of this and other similar adaptive processes.
While the involvement of histone chaperones in the repression of FLO genes is not far fetched, the role of RRM3 calls for a special consideration. It encodes a DNA helicase dispensable for DNA replication over most of the genome, but necessary to relieve pausing at sites of tightly bound proteins . Such pausing sites are frequent in the subtelomeric regions and at the mating type loci , but direct evidence for replication pausing at the FLO genes is not available (Dr. Ivessa, personal communication). It remains possible that the epistatic interactions of RRM3 with CAC1 and ASF1, but not HIR1 (Figs. 1, 4) reflect the susceptibility of the FLO loci to replication pausing and subsequent chromatin perturbation in the absence of chaperones. In this line of thought, both CAF-1 and Rrm3p physically interact with the replication fork clamp PCNA through a conserved PIP (PCNA-Interacting Peptide) motif [39, 40]. Here, we have shown that the flocculation correlated to the ability of CAF-1 to associate with PCNA, thus strongly suggesting that at least the effect of CAF-1 is related to its activity at the replication forks. Our finding may indicate that Rrm3p modulates the association of CAF-1 with PCNA and exacerbates the compromised reassembly of chromatin at paused replication forks . We should also mention that the deletion of RRM3 did not enhance flocculation upon nitrogen starvation or in the presence of NAM (Fig. 5). Further studies will be required to refine the role of Rrm3p in these processes and its precise contribution to chromatin maintenance and to the gene repression.
The role of HIR1 in the repression of FLO genes is also unclear. We have observed apparent flocculation phenotypes in the cac1∆hir1∆ strain and in the hir1∆ strain when exposed to nitrogen starvation or NAM (Figs. 1, 5). On the other hand, the hir1∆rrm3∆ and hir1∆hda1∆ strains showed no phenotype (Fig. 4). It remains possible that the effects of HIR1 deletion are related to the proposed secondary role of HIR in replication-coupled nucleosome assembly in the absence of CAF-I . Alternatively, HIR1 has a limited effect on the repression of FLO genes that can be revealed only after compromising their repression by other means.
Most of the studies on gene repression in budding yeast have focused on SIR-dependent silencing at subtelomeric and mating type loci [8, 41]. At these positions gene silencing is executed by cis-acting silencers that serve as an assembly point of the Sir3/4 proteins, which recruit the Sir2p histone deacetylase and initiate the spreading of histone deacetylation away from the silencers . As already mentioned, FLO genes are silenced by the binding of the Tup1/Cyc8 and Sfl1 repressors upstream of the promoters of these genes and utilize the HDA1, HST1 and RPD3 but not the SIR2 histone deacetylase [4, 7, 42]. It can be said, within the limitations of our current knowledge, that the regulation of the FLO genes and the regulation of gene silencing at the telomeres and the mating loci represent different mechanisms. At the same time, multiple studies have linked SIR-dependent gene silencing to various DNA replication factors and histone chaperones (reviewed in ). Here, we compared the findings in this manuscript to previously published studies (Table 1). We found that the deletions of individual genes or combination of genes show a similar trend of loss of silencing at the mating type loci and sub-telomeres and the loss of repression of the FLO genes. We have also found that treatment of selected mutants with NAM increases flocculation and decreases the silencing at the mating HMR locus (KS, not shown). These correlation analyses support the idea that despite the difference in the mechanisms that establish repression, all these loci share similar requirement for histone chaperones and RRM3 (Table 1).
Another similarity worth mentioning is that subtelomeric genes, partially de-repressed mating loci and at least FLO11 are meta-stable, meaning that they infrequently switch between active and silent state [1, 8]. Here, we have shown that the elevated expression of FLO11 in a flocculating laboratory strain reflects a wide range of the abundance of Flo11p in individual cells. We are not certain if FLO11 alone or all FLO genes variegate. However, it is tempting to speculate that if all FLO genes variegate, they would provide a wide repertoire of cells adhesion patterns, which in turn would aid the adaptation in response to changes in the environment. It is also possible that the varying abundance of GFP as driven by the FLO11 promoter reflects a competition between all FLO genes for regulatory transcription and chromatin factors and that this competition is a key to the variegated expression of these genes. This matter deserves a special attention in future studies.
We show that the deregulation of chromatin transmission and maintenance is sufficient to reconstitute flocculation in laboratory yeast strains. These observations suggest that in these strains lack of flocculation is mediated by epigenetic repression at FLO gene loci. Our paper highlights the FLO genes as attractive loci for future investigation of how epigenetic silencing can drive adaptation and the acquisition of novel phenotypes. Finally, the adverse effects of yeast pathogens like C. albicans and C. glabrata are linked to dimorphic transitions, which involve genes homologous to the FLO genes in S. cerevisiae [1, 43]. Hence, our study might indirectly shed light on the epigenetic basis of this significant health problem.
Materials and methods
The strains used in this study are listed in Additional file 1: Table S1. All assays were conducted with haploid strains in BY4742 and W303 background. Double deletion mutants were produced by routine mating and sporulation. Cells were routinely grown on YPD or SC dropout plates at 30 °C with the exception of temperature-sensitive mutants which were maintained at 23 °C. Liquid cultures were grown on a spinning wheel, not a shaker, to better reveal flocculation. Growth rates of all cultures were measured in ThermoScientific Multiskan G0 instrument. For the nitrogen starvation experiments the strains were grown for 2 days at 30 °C on a spinning wheel in SC medium containing 1% YNB. For the carbon starvation assays SC medium was supplemented with 0.1% glucose and 2% glycerol.
Assessment of flocculation phenotypes
Flocculation was determined by visual observation of cell clusters (Fig. 1) and cell aggregation was confirmed by light microscopy. Sedimentation rates were estimated by resting culture tubes and measuring the time needed for the clearance of the upper 50% of the culture (TS50). The measurement of sedimentation rates in the presence of nicontinamide (NAM) was conducted by dividing the TS50 for the cultures grown in NAM divided by TS50 in the corresponding NAM-free cultures.
RNA was isolated with TRIzol™ solution according to manufacturer’s directions, except that samples were vortexed for 5 min (30 s on, 30 s off) in the presence of equal volume glass beads and precipitated with Ethanol. RNA concentration and purity was determined by ThermoScientific NanoDrop 8000. cDNA synthesis was performed using Applied Biosystems High-capacity cDNA Reverse Transcription Kit. Quantitative PCR was carried out using Applied Biosystems StepOne™Plus thermocycler and PowerUp SYBR Green Master Mix. 6.25 ng of cDNA was added to each reaction and each sample was analyses in triplicates. Quantitative expression values were determined using the ΔΔCq method, wherein the average Cq for each FLO gene was normalised to Cq values for ACT1. Three to five independent experiments were performed with each strain/primer combination and average values, standard deviations and t tests were calculated in Microsoft Excel®. The ΔCq values for each strain/primer combination were normalised to ΔCq values obtained for BY4742 cells and fold expression was calculated as 2−ΔΔCq and shown in a bar graph format.
50 mL cultures were grown to OD600 ~ 0.8, pelleted and washed once in 1 mL of LB buffer (20 mM Tris pH 7. 5 mM EDTA, 140 mM NaCl). Cells were resuspended in 300 μL LB plus 1.5× protease inhibitors (G BioSciences ProteaseArrest™ Yeast/Fungal) and lysed with 500 μL of glass beads for 18 cycles of 30 s ON/30 s OFF with a VWR Pulsing Vortex Mixer. Lysates were spun at 13,000 rpm for 15 min, the supernatant was removed and the pellet resuspended in 500 μL of MNB (200 mM CaCl2, 20 mM Tris pH 7.5, 140 mM NaCl, 1× protease inhibitors). 4 U of MNase was added and the samples were incubated for 5 min at 37 °C followed by the addition of 1/10 volume of STOP (25 mM EDTA, 100 mM EGTA, 140 mM NaCl). Lysates were diluted to 150 μg of DNA in 1.1 mL IP buffer (50 mM Tris pH 7.5, 10 mM EDTA, 140 mM NaCl, 1× protease inhibitors, 0.5% TX-100, 0.15% Deoxycholic acid) and pre-cleared for 1 h with 50 μL of Protein A Sepharose® 4B (Invitrogen.) 250 μL of the pre-cleared lysates were dispensed to tubes containing the relevant antibodies (Millipore 07-352, Milipore 06-866 and Milipore 17-10046) or control antibody (rabbit serum; Millipore 17-10046) and incubated overnight, followed by addition of 40 μL of Protein A Sepharose and a further incubation for 1 h. The beads were washed twice with IP buffer, once with IP buffer plus 360 mM NaCl, once with LiCl buffer (0.25 M LiCl, 1 mM EDTA, 10 mM Tris pH 7.5, 0.5% TX-100) and once TE containing 0.2% TX-100, then resuspended in 100 μL TE with 0.2% TX-100 plus 1 μg of RNase A and incubated at 37 °C for 1 h, then overnight at 65 °C in the presence of 1% SDS followed by 2 h at 37 °C in the presence of Proteinase K. DNA was purified using the GenepHlow™ Gel/PCR kit and eluted into 100 μL of 10 mM Tris. 5 μL of this sample were analysed by qPCR with primers for the ACT1, FLO1 or FLO11 promoters using PowerUp SYBR Green 2× MasterMix (Applied Biosystems) and Applied Biosystems StepOne Plus™ thermocycler with StepOne Software. Three technical replicates were used to calculate a “fold over background” for each of the Histone H3, H3AC and H4AC immunoprecipitations, normalized to ACT1 and then to signals from IPs with rabbit serum, using the formula 2(CtIPserum−CtIP). The relative histone acetylation at each of these positions was determined by normalising the values from the H3, H3AC and H4AC immunoprecipitations to those from Histone 3. The results represent the average of 2 to 3 biological replicas for each strain/gene combination. Average values, standard deviations and t tests were calculated in Microsoft Excel®.
W303 and isogenic cac1∆, rrm3∆ cac1∆rrm3∆ strains were produced by replacement of FLO11 ORF and promoter with a GFP-KanMX cassette as in . All replacements were confirmed by PCR. Cell clusters of these strains were produced by serially diluting cells in 96-well trays and growing them without shaking for 24 h. Images were taken directly from the wells. Cells were also prepared by vigorous vortexing/pipetting of liquid cultures and spreading on slides. All images were taken with a Leica DM 6000B microscope with bright field or with the 469 nm filter. Images were processed and over-layered with Velocity™ software. The quantifying of GFP signals was done by subtracting the background pixel values (ROI with no cells) from the pixel values of identical ROI centred over isolated GFP+ or GFP− cells. For the calculation of the percent of GFP+ cells in Fig. 6b, only cells with 3 times higher signal as compared to the average signal in GFP− cells were counted.
Analysis of FLO gene length variation
DNA from saturated liquid cultures was isolated and subjected to PCR with primers flanking FLO1, FLO5, FLO9, FLO10 and FLO11. Primer sequences were exactly as in . Primer sequences and coordinates in the genome are available upon request.
Analysis of cell cycle
Exponentially growing cultures (OD600 = 1.0) were harvested, fixed in Ethanol and stained with propidium iodine as in . Absorbance was measured by FC500 flow cytometer (Beckman–Coulter) and analysed was performed by FCS express 6 Plus software.
Availability of data and materials
All data generated or analysed during this study are included in this published article and its Additional file.
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We thank Z. Zhang, P. Kaufman, B. Stillman, J. Rine, A. Ivessa, H. Murphy, C. Boone and G.van der Merwe for providing yeast strains; G.van der Merwe and R. Shapiro for valuable discussions.
This study was supported by the National Science and Engineering Research Council of Canada (NSERC) Grant (RGPIN-2015-06727) to KY and financial support to HR and KS by the University of Guelph.
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